The present invention relates to a method of producing a non-human, mammalian oocyte carrying a modified target sequence in its genome, the method comprising the steps of introducing into a non-human, mammalian oocyte: (a) a clustered, regularly interspaced, short palindromic repeats (CRISPR)-associated protein 9 (Cas9 protein) or a nucleic acid molecule encoding said Cas9 protein; and (b-i) a target sequence specific CRISPR RNA (crRNA) and a trans-activating crRNA (tracr RNA) or a nucleic acid molecule encoding said RNAs; or (b-ii) a chimaeric RNA sequence comprising a target sequence specific crRNA and tracrRNA or a nucleic acid molecule encoding said RNA; wherein the Cas9 protein introduced in (a) and the RNA sequence(s) introduced in (b-i) or (b-ii) form a protein/RNA complex that specifically binds to the target sequence and introduces a single or double strand break within the target sequence. The present invention further relates to the method of the invention, wherein the target sequence is modified by homologous recombination with a donor nucleic acid sequence further comprising the step: (c) introducing a nucleic acid molecule into the cell, wherein the nucleic acid molecule comprises the donor nucleic acid sequence and regions homologous to the target sequence. The present invention also relates to a method of producing a non-human mammal carrying a modified target sequence in its genome.
In this specification, a number of documents including patent applications and manufacturer's manuals is cited. The disclosure of these documents, while not considered relevant for the patentability of this invention, is herewith incorporated by reference in its entirety. More specifically, all referenced documents are incorporated by reference to the same extent as if each individual document was specifically and individually indicated to be incorporated by reference.
Gene targeting in embryonic stem (ES) cells is routinely applied to modify the mammalian genome, in particular the mouse genome, which established the mouse as the most commonly used genetic animal model (Capecchi M R (2005)). The basis for reverse mouse genetics was initially established in the 1980-ies, when ES cell lines were established from cultured murine blastocysts, culture conditions were identified that maintain their pluripotent differentiation state in vitro (Evans M J, Kaufman M H., Nature 1981; 292:154-6; Martin G R. Proc Natl Acad Sci USA 1981; 78:7634-8) and it was found that ES cells are able to colonize the germ line in chimaeric mice upon microinjection into blastocysts (Bradley et al., Nature 1984; 309:255-6; Gossler et al., Proc Natl Acad Sci USA 1986; 83:9065-9). Since the first demonstration of homologous recombination in ES cells in 1987 (Thomas K R, Capecchi M R., Cell 1987; 51:503-12) and the establishment of the first knockout mouse strain in 1989 (Schwartzberg P L, Goff S P, Robertson E J., Science 1989; 246:799-803) gene targeting was adopted to a plurality of genes and has been used in the last decades to generate more than 3000 knockout mouse strains that provided a wealth of information on in vivo gene functions (Collins F S, Rossant J, Wurst W., Cell 2007; 128:9-13; Capecchi, M. R., Nat Rev Genet 2005; 6: 507-12). Accordingly, gene targeting in ES cells has revolutionised the in vivo analysis of mammalian gene function using the mouse as genetic model system. However, at present this reverse genetics approach is restricted to mice, as germ line competent ES cell lines that can be genetically modified could be established only from these animals, so far. The exception from this rule is achieved by homologous recombination in primary cells from pig and sheep followed by the transplantation of nuclei from recombined somatic cells into enucleated oocytes (cloning) (Lai L, Prather R S. 2003. Reprod Biol Endocrinol 2003; 1:82; Gong M, Rong Y S. 2003. Curr Opin Genet Dev 13:215-220). However, since this methodology is inefficient and time consuming it did not develop into a simple routine procedure.
Although the generation of targeted mouse mutants as described above is by now well established as a routine procedure, this approach has the drawback that is usually requires a long time of hands on work for vector construction, ES cell culture and selection and the breeding of chimaeras. Additional problems that are often encountered during a gene targeting project are the low efficiency of homologous recombination in ES cells and the loss of the germ line competence of ES cells during the long in vitro culture and selection phase. Therefore, the successful generation of even a single line of knockout mice requires considerable time, the combined efforts of specialists in molecular biology, ES cell culture and embryo manipulation, and the associated technical infrastructure.
Experiments in model systems have demonstrated that the frequency of homologous recombination of a gene targeting vector is strongly increased if a double strand break is induced within its chromosomal target sequence (Rouet, P., Smih, F., Jasin, M.; Mol Cell Biol 1994; 14: 8096-8106; Rouet, P., Smih, F. Jasin, M.; Proc Natl Acad Sci USA 1994; 91: 6064-6068). In the absence of a gene targeting vector for homology directed repair, the cells frequently close the break by non-homologous end-joining (NHEJ). Since this mechanism is error-prone it frequently leads to the deletion or insertion of multiple nucleotides at the cleavage site. If the cleavage site is located within the coding region of a gene it is thereby possible to identify and select mutants that exhibit reading frameshift mutations from a mutagenised population and that represent non-functional knockout alleles of the targeted gene.
Direct genome editing by zinc-finger nucleases (ZFN) as well as TAL-nucleases in one-cell embryos has been recently established as a double strand break-based mutagenesis approach in mice, rats, rabbits and zebrafish (Carbery et al. (2010) Genetics 186:451-9; Cui et al. (2011) Nat Biotechnol 29:64-7; Doyon et al. (2008) Nat Biotechnol 26:702-8; Flisikowska et al. (2011) PLoS One 6:e21045; Meyer et al. (2010) Proc Natl Acad Sci USA 107:15022-6; Geurts A M, et al. (2009) Science 325:433; Huang (2011) Nat Biotechnol 29:699-700; Tesson (2011) Nat Biotechnol 29:695-696). Such nucleases are designed to induce double-strand breaks (DSBs) at preselected genomic target sites (Klug (2010) Annu Rev Biochem 79:213-231; Porteus & Carroll (2005) Nat Biotechnol 23:967-73; Porteus & Baltimore (2003) Science 300:763; Santiago et al. (2008) Proc Natl Acad Sci USA 105:5809-14). DSBs targeted to coding exons frequently undergo sequence deletions leading to gene knockout or allow the insertion (knock-in) of DNA sequences from gene targeting vectors via homologous recombination (HR). The generation of knockout and knock-in mutants at the Rosa26, Mdr1a, Pxr, and IgM loci by microinjection of ZFNs one-cell embryos of mice, rats and rabbits (Cui et al. (2011) Nat Biotechnol 29:64-7; Flisikowska et al. (2011) PLoS One 6:e21045; Meyer et al. (2010) Proc Natl Acad Sci USA 107:15022-6; Huang (2011) Nat Biotechnol 29:699-700; Tesson (2011) Nat Biotechnol 29:695-696) has recently been reported.
In addition, TAL elements have been combined with the Fokl nuclease domain to create TAL-nuclease fusion proteins (TALENs) that enable to generate double-strand breaks within intended target regions (Christian M et al. (2010). Genetics 186:757-761; Cermak et al. (2011) Nucleic Acids Res 39:e82; Miller et al. (2011) Nat Biotechnol 29:143-148). TALENs were shown to enable gene editing in mammalian cell lines and in zebrafish, mouse and rat embryos (Sung et al. (2013) Nat Biotechnol 31:23-24; Tesson et al. (2011). Nat Biotechnol 29:695-696; Reyon et al. (2012). Nat Biotechnol 30:460-465).
However, even though the use of zinc finger nucleases results in a higher frequency of homologous recombination, considerable efforts and time are required to design zinc finger proteins that bind a new DNA target sequence at high efficiency. In addition, it has been calculated that using the presently available resources only one zinc finger nuclease could be found within a target region of 1000 basepairs of the mammalian genome (Maeder, et al. 2008 Mol Cell 31(2): 294-301; Maeder, et al. 2009 Nat Protoc 4(10): 1471-501). Further, the use of TALENs involves considerable efforts since it requires the de novo construction and expression of two large TAL-nuclease fusion proteins specifically for each target site. Also, the principles of the TAL peptide DNA recognition are still not fully understood, thus often leading to the necessity of time- and cost-consuming further experimentations in order to optimize the respective TALENs.
Recently, a novel system for inducing single or double strand breaks in target nucleic acid sequences has been found. This system is referred to in the art as CRISPR/Cas system, which stands for “clustered, regularly interspaced, short palindromic repeats (CRISPR)/CRISPR-associated protein”. It is based on an adaptive defense mechanism evolved by bacteria and archaea to protect them from invading viruses and plasmids, which relies on small RNAs for sequence-specific detection and silencing of foreign nucleic acids. CRISPR/Cas systems are composed of cas genes organized in operon(s) and CRISPR array(s) consisting of genome-targeting sequences (called spacers) interspersed with identical repeats (Bhaya et al. (2011) Annu Rev Genet 45:273-297; Barrangou R, Horvath P (2012) Annu Rev Food Sci Technol 3:143-162). CRISPR/Cas-mediated immunity in bacteria and archaea occurs in three steps. In the adaptive phase, bacteria and archaea harboring one or more CRISPR loci respond to viral or plasmid challenge by integrating short fragments of foreign sequence (protospacers) into the host chromosome at the proximal end of the CRISPR array. In the expression and interference phases, transcription of the repeat spacer element into precursor CRISPR RNA (pre-crRNA) molecules followed by enzymatic cleavage yields short crRNAs (CRISPR RNAs) that can subsequently pair with complementary protospacer sequences of invading viral or plasmid targets. Target recognition by crRNAs directs the silencing of the foreign sequences by means of Cas proteins that function in complex with the crRNAs.
There are three types of CRISPR/Cas systems (Makarova et al. (2011) Nat Rev Microbiol 9:467-477). The type I and III systems share some overarching features: specialized Cas endonucleases process the pre-crRNAs, and once mature, each crRNA assembles into a large multi-Cas protein complex capable of recognizing and cleaving nucleic acids complementary to the crRNA.
In contrast, type II systems process precrRNAs by a different mechanism in which a trans-activating crRNA (tracrRNA) complementary to the repeat sequences in pre-crRNA triggers processing by the double-stranded RNA specific ribonuclease RNase III in the presence of Cas9 (formerly Csn1) protein. Cas9 is the sole protein responsible for crRNA-guided silencing of foreign DNA.
Jinek et al. recently demonstrated that the Cas9 endonuclease family can also be programmed with single “chimaeric” RNA molecules, containing a target recognition sequence at the 5′ end followed by a hairpin structure retaining the base-pairing interactions that occur between the tracrRNA and the crRNA (Jinek et al. (2012 Science 337:816-821). This single transcript effectively fuses the 3′ end of crRNA to the 5′ end of tracrRNA, thereby mimicking the dual-RNA structure required to guide site-specific DNA cleavage by Cas9.
The Streptococcus pyogenes SF370 type II CRISPR locus consists of four genes, including the Cas9 nuclease, as well as two non-coding RNAs: tracrRNA and a pre-crRNA array containing nuclease guide sequences (spacers) interspaced by identical direct repeats (DRs) (Deltcheva et al. (2011) Nature 471:602-607).
Cong et al. (Cong et al. (2013). Science 339:819-823) recently applied this prokaryotic RNA-programmable nuclease system to introduce targeted double stranded breaks (DSBs) in mammalian chromosomes through heterologous expression of the key components. It has been previously shown (Jinek et al. (2012 Science 337:816-821) that expression of tracrRNA, pre-crRNA, host factor RNase III, and Cas9 nuclease are necessary and sufficient for cleavage of DNA in vitro. Expression of a codon optimized S. pyogenes Cas9 (SpCas9), of an 89-nucleotide (nt) tracrRNA and of a pre-crRNA comprising a single guide spacer flanked by DRs was expressed in human 293 cells. The initial spacer was designed to target a 30-basepair (bp) site (protospacer) in the human EMX1 locus that precedes an NGG, the requisite protospacer adjacent motif (PAM). Heterologous expression of the CRISPR system (SpCas9, SpRNase III, tracrRNA, and pre-crRNA) achieved targeted cleavage of mammalian chromosomes. In addition, a chimeric crRNA-tracrRNA hybrid was used, where a mature crRNA is fused to a partial tracrRNA via a synthetic stem-loop to mimic the natural crRNA:tracrRNA duplex. Cong et al. observed cleavage of all protospacer targets when SpCas9 was co-expressed with pre-crRNA (DRspacer-DR) and tracrRNA. Furthermore, Cong et al. showed that also the Streptococcus thermophilus LMD-9 CRISPR1 system can mediate mammalian genome cleavage.
In another recent report, Mali et al. (Mali et al. (2013); Science 339: 823-826) independently confirmed high efficiency CRISPR-mediated genome targeting in several human cell lines, while Hwang et al. (Hwang et al. (2013); Nature Biotechnology doi:10.1038/nbt.2501) showed that this system may also be employed in zebrafish.
Whereas this system has been shown to be functional in mammalian cells such as human embryonal kidney cells (such as e.g. 293T or 293 FT cells), human chronic myeloid leukemia cells (such as K562 cells) or induced pluripotent stem cells, no attempts have been reported to employ this system in oocytes/zygotes.
As totipotent single entities, mammalian zygotes could be regarded as a preferred substrate for genome engineering since the germ line of the entire animal is accessible within a single cell. However, the experimental accessibility and manipulation of zygotes is severely restricted by the very limited numbers at which they are available (dozens-hundred) and their very short lasting nature. These parameters readily explain that the vast majority of genome manipulations, that occur at frequencies of below 10−5 like gene targeting, can be successfully performed only in cultured embryonic stem cells that are grown up to a number of 107 cells in a single standard culture plate. The only exception from this rule concerns the generation of transgenic mice by pronuclear DNA injection that has been developed into a routine procedure due to the high frequency of transgene integration in up to 30% of injected zygotes (Palmiter R D, Brinster R L.; Annu Rev Genet 1986; 20:465-499). Since microinjected transgenes randomly integrate into the genome, this method can only be used to express additional genes on the background of an otherwise normal genome, but does not allow the targeted modification of endogenous genes.
An early report to characterize the potential of zygotes for targeted gene manipulation by Brinster (Brinster R L, Braun R E, Lo D, Avarbock M R, Oram F, Palmiter R D.; Proc Natl Acad Sci USA 1989; 86:7087-7091) showed that this approach is not practical as only one targeted mouse was obtained from >10.000 zygotes within 14 months of injections. Thus, Brinster et al. discouraged any further attempts in this direction. In addition to a low recombination frequency, Brinster et al. noted a high number of spontaneously occurring, undesired mutations within the targeted allele that severely compromised the function of the (repaired) histocompatibility class II gene. From the experience of Brinster et al. it could be extrapolated that the physiological, biochemical and epigenetic context of genomic DNA in the zygotic pronuclei are unfavourable to achieve targeted genetic manipulations, except for the random integration of transgenes that occurs at high frequency.
In addition, the biology of oocyte development into an embryo provides further obstacles for targeted genetic manipulations.
A growing mouse oocyte, arrested at diplotene of its first meiotic prophase, transcribes and translates many of its own genes, thereby producing a store of proteins sufficient to support development up to the 8-cell stage. These transcripts guide oocytes on the two steps of oocyte maturation and egg activation to become zygotes. Typically, oocytes are ovulated and become competent for fertilisation before reaching a second arrest point. When an oocyte matures into an egg, it arrests in metaphase of its second meiotic division where transcription stops and translation of mRNA is reduced. At this point an ovulated mouse egg has a diameter of 0.085 mm and, with a volume of ˜300 picoliter, it exceeds the size of a typical somatic cell by a 1000-fold (Nagy A, Gertsenstein M, Vintersten K, Behringer R., 2003. Manipulating the Mouse Embryo. Cold Spring Harbour, N.Y.: Cold Spring Harbour Laboratory Press). The re-modeling of a fertilised oocyte into a totipotent zygote is one of the most complex cell transformations in biology. Remarkably, and in stark contrast to other mammalian cell types, this transition occurs in the absence of transcription factors and therefore depends on proteins and mRNAs accumulated in the oocyte during oogenesis. The embryonic development of a mammal begins when sperm fertilises an egg to form a zygote. Fertilization of the egg triggers egg activation to complete the transformation to a zygote by signaling the completion of meiosis and the formation of pronuclei. At this stage the zygote represents a 1-cell embryo that contains a haploid paternal pronucleus derived from the sperm and a haploid maternal pronucleus derived from the oocyte. In mice this totipotent single cell stage lasts for only ˜18 hours until the first mitotic division occurs.
In fertilized mammalian eggs, the two pronuclei that undergo DNA replication, do not fuse directly but approach each other and remain distinct until the membrane of each pronucleus has broken down in preparation for the zygote's first mitotic division that produces a 2-cell embryo. The 1-cell zygote stage is characterised by unique transcriptional and translation control mechanisms. One of the most striking features is a time-dependent mechanism, referred to as the zygotic clock, that delays the expression of the zygotic genome for ˜24 h after fertilization, regardless of whether or not the one-cell embryo has completed S phase and formed a two-cell embryo (Nothias J Y, Majumder S, Kaneko K J, DePamphilis M L.; J Biol Chem 1995; 270:22077-22080). In nature, the zygotic clock provides the advantage of delaying zygotic gene activation (ZGA) until chromatin can be remodelled from a condensed meiotic state to one in which selected genes can be transcribed. Since the paternal genome is completely packaged with protamines that must be replaced with histones, some genes would be prematurely expressed if ZGA were not prevented. Cell-specific transcription requires that newly minted zygotic chromosomes repress most, if not all, promoters until development progresses to a stage where specific promoters can be activated by specific enhancers or trans-activators. In the mouse, formation of a 2-cell embryo marks the transition from maternal gene dependence to zygotic gene activation (ZGA). Among mammals, the extent of development prior to zygotic gene activation (ZGA) varies among species from one to four cleavage events. Maternal mRNA degradation is triggered by meiotic maturation and 90% completed in 2-cell embryos, although maternal protein synthesis continues into the 8-cell stage. In addition to transcriptional control, the zygotic clock delays the translation of nascent mRNA until the 2-cell stage (Nothias J Y, Miranda M, DePamphilis M L.; EMBO J 1996; 15:5715-5725). Therefore, the production of proteins from transgenic expression vectors injected into pronuclei is not achieved until 10-12 hours after the appearance of mRNA.
WO2011/051390 describes a method for modifying a target sequence in the genome of a mammalian or avian oocyte by homolgous recombination using a zinc finger nuclease and, thus, a method of producing a non-human mammal carrying a modified target sequence in its genome. However, since this method makes use of a zinc finger protein, it is associated with the drawbacks described above with regard to zinc finger proteins. No indication is provided in WO2011/051390 that successful recombination in oocytes could be achieved by any other means but zinc finger proteins.
WO2011/154393 describes a method of modifying a target sequence in the genome of a eukaryotic cell, wherein a fusion protein comprising a DNA-binding domain of a Tal effector protein and a non-specific cleavage domain of a restriction nuclease is employed to introduce a double strand break within the target sequence, thereby enhancing the modification of the target sequence by homologous recombination. It is further described that the method can be applied to oocytes and that it can be used to produce a non-human mammal or vertebrate carrying a modified target sequence in its genome. However, the only methods for introducing double strand breaks and enhancing the frequency of homologous recombination that are described in WO2011/154393 are the use of zinc finger proteins or fusion proteins comprising a DNA-binding domain of a Tal effector protein and a non-specific cleavage domain of a restriction nuclease. No reference is made to the CRISPR/Cas system and no indication is provided that the frequency of homologous recombination in oocytes could be enhanced by any means other than Zinc finger proteins or the claimed fusion proteins.
Thus, whereas methods have been described in the art for the generation of transgenic animals carrying targeted modifications in their genome, there is still a need to provide means to generate genetically modified animals faster, easier and more cost-effective than using any of the prior art methods.
This need is addressed by providing the embodiments characterized in the claims.